Helium Ion Microscopy (HIM) for theimaging of biological samples atsub-nanometer resolutionMatthew S. Joens1, Chuong Huynh2, James M. Kasuboski1, David Ferranti2, Yury J. Sigal1,Fabian Zeitvogel3, Martin Obst3, Claus J. Burkhardt4, Kevin P. Curran5, Sreekanth H. Chalasani5,Lewis A. Stern2, Bernhard Goetze2 & James A. J. Fitzpatrick1

1Waitt Advanced Biophotonics Center, Salk Institute for Biological Studies, 10010 North Torrey Pines Road, La Jolla, CA 92037,USA, 2Ion Microscopy Innovation Center, Carl Zeiss Microscopy LLC, One Corporation Way, Peabody, MA 01960, USA, 3Centerfor Applied Geosciences, University Tubingen, Hoelderlinstr. 12, 72074 Tuebingen, Germany, 4NMI Natural and MedicalSciences Institute, Markwiesenstr. 55, 72770 Reutlingen, Germany, 5Molecular Neurobiology Laboratory, Salk Institute forBiological Studies, 10010 North Torrey Pines Road, La Jolla, CA 92037, USA.

Scanning Electron Microscopy (SEM) has long been the standard in imaging the sub-micrometer surfaceultrastructure of both hard and soft materials. In the case of biological samples, it has provided great insightsinto their physical architecture. However, three of the fundamental challenges in the SEM imaging of softmaterials are that of limited imaging resolution at high magnification, charging caused by the insulatingproperties of most biological samples and the loss of subtle surface features by heavy metal coating. Thesechallenges have recently been overcome with the development of the Helium Ion Microscope (HIM), whichboasts advances in charge reduction, minimized sample damage, high surface contrast without the need formetal coating, increased depth of field, and 5 angstrom imaging resolution. We demonstrate the advantagesof HIM for imaging biological surfaces as well as compare and contrast the effects of sample preparationtechniques and their consequences on sub-nanometer ultrastructure.

The Scanning Electron Microscope (SEM) has become one of the fundamental tools used in the study ofexterior morphology and structure for biological samples1. More recently it has become an essential tool inthe study of connectomics and cellular interactions with advances in serial block-face imaging techniques,

for example, using either an in situ microtome2 or focused-ion beam (FIB) milling3. The caveat, however, is thatconventional SEM imaging requires samples to be conductive and vacuum friendly, both of which can beaccomplished via drying of the sample and coating it with a heavy metal1. Although this method of samplepreparation has been standard for many years, there are some instances where minute surface details can beobscured by the metal coating or where the sample is too delicate or small to survive the requisite fixation anddrying procedures4–6. To combat this, numerous advances have been made to reduce charge as well as imagesamples in their hydrated form.

One of the most effective methods used to minimize the charging effects observed in insulating samples is toreduce the accelerating voltage of the electron beam, also known as low-voltage scanning electron microscopy(LVSEM)7. Lowering the voltage in conjunction with brighter field-emission sources and state of the art detectorshas the advantage of higher resolution, increased topographical contrast and surface sensitivity as well asincreased secondary electron generation. However, as the voltage decreases, chromatic aberration increases,and depth of field decreases with the necessary decrease in working distance, thus limiting the low voltage rangeof most conventional non aberration-corrected SEMs8–10. An alternative approach is to delay the dehydration ofsamples using variable and extended pressure imaging techniques11,12. By injecting nitrogen gas (variable pres-sure) or water vapor (extended pressure) into the imaging chamber, charge can be negated by creating secondaryionization products of the working gas molecules1. Beam stability is maintained in these systems by means of anIn-Lens differential pumping system that allows the maintenance of a high vacuum in the electron gun with a lowvacuum in the imaging chamber. Although useful, this technique is limited in resolution and signal to noise as aresult of beam skirting13. In addition, there is an increased chance of beam-induced damage to the sample due tothe higher accelerating voltages required for imaging within a gaseous environment14.

OPEN

SUBJECT AREAS:IMAGING

TECHNIQUES ANDINSTRUMENTATION

Received2 October 2013

Accepted26 November 2013

Published17 December 2013

Correspondence andrequests for materials

should be addressed toJ.A.J.F. (fitzp@salk.

edu)

SCIENTIFIC REPORTS | 3 : 3514 | DOI: 10.1038/srep03514 1

One of the most formidable developments in imaging biologicalsurfaces in their most native form without any chemical fixation isCryo-SEM15, which freezes or cryo-immobilizes the sample andtransfers it to a chilled stage prior to imaging. Although it is possibleto maintain gross morphology with this technique, samples canpotentially sublimate under high vacuum conditions thus increasingthe risk for charging. This often requires the frozen samples to besputter coated or imaged in a variable pressure environment14,16.

Notwithstanding these advances in controlling charge and main-taining biological surface structure, they all still suffer from limita-tions in both resolution and contrast. Recently, an ultra-highbrightness gas field ionization source (GFIS) has been developedfor use as an ultra-high resolution and high contrast imaging system(Orion, Carl Zeiss, Peabody, MA)17. Despite this relatively contem-porary innovation, the original concept of a Field Ion Microscope(FIM) based on helium ions was originally proposed by Erwin Mullerin 195618. However, recent refinements to the design overcame pre-vious limitations and now boast a reproducible and stable ion sourceover many days. The key to the systems performance is its source,which is shaped to a three-sided pyramidal tip consisting of onlythree atoms at the apex. When the tip is cryogenically cooled andexposed to a high voltage (30 kV) in the presence of helium, anionized gas field is generated at the apex of the tip. This is due tofield only being intense enough at those three atoms to efficientlyionize the helium gas. As such, the ion emission originates preferen-tially from those atoms at the apex, of which only one is selected forimaging. Thus imaging occurs using a single atom as the ion source.The extracted ions are accelerated down the column of the micro-scope in the form of a beam that is then shaped and raster scannedmuch in the same fashion as in an SEM. Due to the ultra highbrightness of the GFIS a very small beam defining aperture may beused and therefore spherical and chromatic aberrations of the ionoptical column are insignificant. The helium ion microscope there-fore exhibits substantially higher resolution with a larger depth offield resulting in an estimated resolution that is five times greaterthan that of a modern FE-SEM17. Also, since the de Broglie wave-length of a helium ion is very small (below one picometer for a 30 kVaccelerating voltage) the spot size of the scanned beam is not limitedby diffraction aberration17. In addition to the increase in resolution,the increased mass of a helium ion results in a strong forward scat-tering of the ions causing the majority of the beam energy to bedeposited in deeper areas of sample. This results in high contrastsecondary electron emission deriving mostly from the primary beam(SE1 electrons), with typically 2–5 electrons produced per heliumion19. Because of the deeper penetration of the forward scattered ionsin biological samples, there is a much lower propensity for beam-induced surface-damage due to lower sample densities as comparedwith crystalline materials such as silicon. The lack of deposition ofenergy at the sample surface results in minimal damage at 30 kVwhere currents used for imaging range from 0.1–0.4 pA19–22.Furthermore, the sputter rate of helium is two orders of magnitudelower than gallium at comparable beam energies. This results inbiological samples being able to be imaged repetitively for a relativelylong time before any noticeable beam damage occurs. Such effectsmay be visible as slight deformations or almost negligible materialremoval (a polishing effect) from the sample surface. For electricallyinsulating samples, positive charge resulting from the ion beam iseasily compensated by directing a very low voltage electron beam(flood gun) at the sample. The use of such a device does not have anyaffect on the helium ion spot size and thus has no negative impact onresolution19. Also, the recent development of patterning scan engines(Fibics, Ottawa, Canada) allow the routine nanoscale milling andmodification of samples17.

To date, helium ion microscopy has been primarily been used inhigh-contrast imaging of inorganic materials; however23,24, therehave been some limited instances where it has been used in imaging

of biological samples. These have included imaging nanofibers inhyaluronan-based pericellular matrix25, edge detection of colon can-cer cells26, visualization of articular cartilage networks27 and theultrastructure of Lepidoptera scales28. More recently, HIM has beenused successfully to visualize the microvilli and glomerular structuresin the rat kidney29 and nanoscale cuticle structures on Drosophilamelanogaster30. In addition to using slow helium ions to image,research has been undertaken on the development of a MegaElectron Volt (MeV) microscope for whole cell density mapping31.

Despite these isolated demonstrations of HIM imaging, there hasyet to be any significant interest demonstrated in developing HIMtechnology for biological applications. As such, the aim of this studywas to undertake a set of experiments to test if the advantagesdescribed above would lead to improved insight on biological speci-mens. Therefore the HIM was tested on a traditionally difficult rangeof challenging biological samples. In addition, besides demonstratingthe resolution and contrast abilities of the method, we also comparedresults to a modern FE-SEM as well as identify any potential con-sequences of sample preparation (critical point drying and sputtercoating), as seen in the ultrastructure of the various specimens.

ResultsTo compare SEM to HIM, a set of biological samples were chosenwhere the unique aspects of the HIM system could provide advantagesin their imaging. (i) Arabidopsis thaliana, the most favored modelsystem for plant biology to study exterior nanoscale morphology (ii)HeLa cells, as an adherent cell type to study membrane surface detailsand filopodia, (iii) BoFeN1 iron-oxidizing bacteria to showcase theeffects of drying and heavy metal coating on surface morphology and(iv) Pristionchus pacificus predator nematodes to illustrate the depthof field of the HIM system and its precision milling abilities.

Imaging parameters were chosen to provide a rational comparisonbetween the two technologies. Since the overarching goal of the studywas to investigate uncoated samples as well as to achieve high-reso-lution images at high magnifications, the FE-SEM was operated at alow voltage (0.7–1.0 kV for uncoated samples, and 3 kV for coatedsamples) with an aperture size of 30 mm. Both In-Lens (I-L) andEverhart-Thornley (E-T) detectors were used to image the samplesas only secondary electrons were studied in the HIM. Alternatively,the HIM was operated at an optimal imaging voltage of 30 kV, anaperture size of 5 mm and a beam blanker current of 0.5 pA. AnEverhart-Thornley (E-T) detector was used to image the samples.In both instances, working distance and tilt were slightly variedbetween images and samples to produce optimal imaging conditionsof electron detection, depth of field, and charge compensation.

Care was taken during sample preparation to ensure that anyartifacts were minimal. To reduce drying artifacts and to make thesamples vacuum stable, proper fixation and critical point drying(CPD) was necessary. It was found that small plants and nematodescould be fixed with ice-cold ethanol and then directly critical pointdried. Cells, however, were more sensitive to the CPD process andrequired the use of robust fixatives such as glutaraldehyde to main-tain intact membrane structures. In addition all samples were kept ina desiccator or under vacuum at all times to minimize artifacts causedby rehydration of the tissues from native humidity.

Arabidopsis thaliana. As a model plant, Arabidopsis was chosen as astable and well-studied organism to compare the effects of imaging atever increasing magnification ranges between the FE-SEM and HIM.Since the samples were critical point dried and uncoated, they wereprone to charging and thus low voltage (,1 kV) imaging in the FE-SEM was required. When compared to the HIM, images in the SEMwere similar at lower magnifications in both contrast, depth of field,and lack of charging (Figure 1a and b panels). However, despite low-voltage imaging in the FE-SEM, the sample exhibited charging effectsstarting at ,20 kX magnification (3.4 mm field of view), resulting in

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motion shown by streaks or bands within the image (Figure 1c, whitearrowheads). In addition, at ,38 kX (2.1 mm field of view) finetextures on the sample surface and minute ridges on the cuticlecould no longer be discerned. By 163 kX (478 nm field of view),the cuticle could no longer be clearly resolved in the FE-SEM as aresult of either heavy charging or beam-induced thermal drift due tothe very small probe size necessary to achieve such a magnification(Figure 1c far right panel). In contrast, charging in the HIM wascompletely eradicated via the use of the microscope’s charge-compensation flood gun. As a result, higher magnifications up to163 kX could be easily achieved without any form of charging,thermal drift and most importantly beam-induced damage, thusrevealing fine surface structures, such as the defects in the wax ona single ridge of the plant’s cuticle (Figure 1d panels).

HeLa cells. As a model cell line used in many cell biology applica-tions, HeLa cells were chosen to demonstrate the effects of imagingfine structures such as filopodia and cell adhesion points in bothHIM and SEM. The comparison of two cells in mitosis at low magni-fication (Figure 2a) clearly shows the efficient charge neutralizationability of the HIM over the FE-SEM. The intensity-saturated regionsof the cell as well as the appearance of dark bands on the glasscoverslip indicated the areas of charging in the FE-SEM. As themagnification was increased to 38 kX (3 mm field of view), twoseparate effects can be seen within each microscope. In the HIM,

filopodia had a tendency to appear ‘‘ghosted’’ or somewhat translu-cent when there were additional features in close z-proximity(Figure 2b, right panel, black arrowheads). As described previ-ously, this is likely due to the deeper penetration of the heliumions that pass through thinner regions of the sample, generatingnot only SE1 electrons on the surface closest to the source, but alsosecondary electrons on the surfaces immediately behind them31. Thisis due to the ion beam not changing geometry over that distance. Inthe SEM, the filopodia exhibited edge effects, a common artifact oflow-voltage imaging of fine or filamentous structures (Figure 2b, leftpanel, black arrowheads). As magnification was increased a thirdtime to 163 kX (700 nm field of view) (Figure 2c), the filopodiawere easily recognizable in both microscopes, however, the surfacefeatures and textures were indistinguishable in the FE-SEM(Figure 2c, left panel).

In addition to the comparison of the HIM to the SEM, the max-imum overall resolution of the HIM was demonstrated in Figure 3. Arandom HeLa cell was chosen (Figure 3a) and from there, a singlefilopodia attachment point was recorded at increasing magnifica-tions (Figure 3b–d) until a maximum magnification of 285 kX, ora field of view of 400 nm was achieved. At this magnification, thesample contrast between the cell and the glass coverslip was still highenough to discern the size of the attachment point (,5 nm).Remarkably, repeated imaging of these small and highly delicatestructures caused no discernable beam damage.

Figure 1 | Comparison of HIM and FE-SEM imaging in Arabidopsis thaliana. (a) Low and (c) high magnification series of uncoated and critical point

dried Arabidopsis thaliana sepal cuticle structures in the FE-SEM at low voltage (,1 kV) using an Everhart-Thornley (E-T) detector. At 22 kX

magnification, charging artifacts become visible in the form of streaks or bands (white arrowheads) that go horizontally across the image. (b) Low and (d)

high magnification series of the sepal cuticle structure in the HIM. Throughout the magnification range, no charging or imaging artifacts are apparent and

cuticle structures are still visible at 163 kX. Scale Bars: (a,b) 10 mm (c,d) 200 nm.

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Iron-oxidizing bacteria (BoFeN1). The nitrate-reducing Acidovo-rax sp. strain BoFeN1, originally isolated from anoxic freshwatersediments of Lake Constance causes oxidation of Fe(II) either byan enzymatic process or by an abiotic reaction of Fe(II) withnitrite, which is an intermediate denitrification product32. It is

known from numerous previous studies that, when grown in thepresence of Fe(II), some cells tend to grow a crust of platelet-shaped Fe(III) minerals on the cell surface while others do not33.

The HIM images that were acquired at low magnification showthree-dimensional cell mineral aggregates (Figure 4c and d, left

Figure 2 | Comparison of HIM and FE-SEM imaging of mitotic HeLa cells. (a) Low magnification of uncoated critical point dried HeLa cells grown on

glass coverslips in metaphase in both the FE-SEM at low voltage (,1 kV) with an In-Lens (I-L) detector (left panel) and HIM (right panel). Whilst the

HIM exhibits high contrast and depth of field, the FE-SEM shows signs of charging in the form of saturated areas and dark and light banding on the

glass coverslip. (b) At ,38 kX (3 mm field of view), the ‘‘ghosting’’ effect of some filopodia becomes apparent in the HIM (right panel, black arrowheads),

while the FE-SEM exhibits a large amount of edge effects (left panel, black arrowheads). (c) High magnification of the same dividing cells shows no

sign of loss of resolution in the HIM compared to the FE-SEM as depicted by the presence of cell membrane textures. Scale Bars: (a) 5 mm (b) 500 nm (c)

100 nm.

Figure 3 | Single filopodia attachment point of a HeLa cell as illustrated using the HIM. (a) A single cell was chosen that had minimal contact with

neighboring cells and visible attachment points to the glass substrate. (b,c) The magnification was increased on a single adhesion point to a final

magnification of (d) ,285 kX (400 nm field of view) while maintaining high enough contrast to depict an attachment width of ,5 nm. Scale Bars: (a)

5 mm (b) 1 mm (c,d) 50 nm.

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SCIENTIFIC REPORTS | 3 : 3514 | DOI: 10.1038/srep03514 4

panels). The images show a large depth of field in comparison to FE-SEM images that were acquired at low voltage at similar magnifica-tions (Figure 4a and b, left panels). Low voltages were necessary inparticular for imaging the uncoated samples by FE-SEM to avoidcharging artifacts. The comparison of coated and uncoated samplesby HIM at high resolution reveals two major artifacts caused by thesputter coating with platinum. Comparing the coated and non-coated critical point dried samples (Figure 4c and d) it appears thatthe crystallites attached to the cells end in rather sharp tips withoutcoating, whereas mineral platelets on the cell surfaces of the plat-inum-coated samples appear to be relatively thick at the end andsometimes even show globule-like structures in the size-range of tensof nanometers (Figure 4c, right panel). A potential explanation forthis change in mineral shape and structure might be a localized,preferential deposition of platinum due to charge-effects on the elec-trically isolating mineral structures. Additionally, the critical pointdried samples show especially clean surfaces, whereas the surfaces ofthe minerals in plunge-frozen and freeze-dried samples appear tohave a much smoother envelope in the case of the uncoated samples.We attribute this to a thin layer of extracellular polymeric substances(EPS) coating the whole aggregate. In contrast, we observed a rela-tively rough surface in the case of the platinum-coated samples (seeSupplementary Fig. 1). This indicates that sputter coating with plat-inum introduces an artificial surface roughness.

HIM imaging has, for the first time, allowed the identification ofthis coating artifact because of the higher spatial resolution in com-bination with the capability of analyzing non-conductive, uncoatedsamples. In addition, the existence of an EPS-envelope coating theminerals in plunge-frozen and freeze-dried samples could only beunambiguously shown in non-coated samples. Furthermore, sputtercoating with platinum seemed only to affect the surface of the chem-ically fixed and critical point dried bacteria by creating a texturedsurface with network-like structures that created high secondaryelectron-signals (Figure 4c). We did not observe this effect whenthe cells were not coated with platinum (Figure 4d). We also didnot observe these structures on plunge-frozen and freeze-dried sam-ples, both with and without the heavy metal coating (see Supple-mentary Fig. 1). Since dehydration in solvents such as ethanol (anecessary step preceding critical point drying), can cause shrinkagein biological samples34, it is also possible that a similar effect couldlead to a slightly textured cell surface that is potentially electricallyinsulating and thus, could favor localized deposition of platinum dueto charge induced effects.

Predator nematode - pristionchus pacificus. Finally the predatornematode, Pristionchus pacificus35 served as an ultrastructural imag-ing challenge in the sense that the interior of the mouth cavity,

including teeth morphology, had never been successfully imaged.A sister nematode, Parasitodiplogaster laevigata, a parasite of figwasps has previously been shown via SEM imaging to have aprotruding tooth structure and a visible Dorsal Esophogeal GlandOrifice (DEGO) located at the base of the protruding tooth36.However, in the initial HIM imaging of Pristionchus pacificus, itwas revealed that a membranous sheath obscured the primarytooth structure (Figure 5a). This afforded an opportunity toemploy another unique aspect of the HIM system – precisionnano-machining using focused noble gas ions. By switching outthe working gas of the HIM from helium to neon, it was possible

Figure 4 | Comparison of platinum coated and uncoated sample preparations critical point dried BoFeN1 bacteria in both the HIM and FE-SEM at lowand high magnifications. (a) Low and high magnifications of platinum coated (using an Everhart-Thornley (E-T) detector) and (b) uncoated bacterium

(using an In-Lens (I-L) detector) in the FE-SEM reveal Fe(III) platelet structures that were on the order of tens of nanometers in size. Although the

structures appeared much more flat in the uncoated sample, resolution was limited due to the necessity to use low-voltage imaging. (c) Low and high

magnifications of platinum coated and (d) uncoated bacterium in the HIM reveal the coating induced artifact of globular structures and artificial

textures on the platelets. The uncoated platelets in the HIM were for the first time revealed to be relatively flat with relatively sharp-tipped cellular

attachments. Scale bars: 250 nm.

Figure 5 | HIM imaging of Pristionchus pacificus. (a) Native imaging of

the nematode revealed the interior mouth cavity and tooth structures were

obscured by a sheath-like structure. (b) neon milling of the outer sheath in

the HIM revealed the primary tooth and DEGO structure of the nematode

with minimal thermal damage to the cutting surface. (c) The removal of

the outer sheath was performed in a very controlled manner with a final

dose of 30 nC/mm3. (d) The delicate nature of the neon milling was

demonstrated by the patterning of a small horizontal line using a dose of

0.3 nC/mm3. Despite being strong enough to penetrate the outer cuticle of

the nematode, minimal ablation and thermal damage is visible on the

surrounding area. Scale Bars: (a–c) 5 mm (d) 1 mm.

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to delicately remove the tip of the membranous sheath, while leavingthe interior of the mouth cavity intact (Figure 5c). This reveals theinterior mouth structure as well as the primary tooth morphologyand DEGO shape and size (Figure 5b). The very precise millingability of the neon beam was effectively demonstrated by cuttingthe exterior of the nematode with a final dose of 0.3 nC/mm3

(Figure 5d). This proved intense enough to cut through the outerskin of the worm without damaging any of the surrounding tissue.For deeper cuts (such as the one shown in Figure 5c), the final cuttingdose was increased to 30 nC/mm3 to penetrate the entire depth of theworm and efficiently mill through some of the tougher, morefilamentous portions of the sample such as the mechanosensoryfibers around the nematode head.

DiscussionOur results demonstrate both novel and exciting applications forhelium ion microscopy in visualizing the surface ultrastructure ofbiological specimens at an unparalleled level of resolution and con-trast. Specifically, we show that the common charging artifacts withthe FE-SEM can be greatly reduced using the HIM approach.Moreover, the low sputtering rate of the helium ions beam fororganic materials enables the repeated imaging of small and delicatesurface features with no discernable beam damage. Also, we showthat the heavy metal coating as generally required for SEM intro-duces clearly resolvable structural artifacts and HIM is a superiormethod in clearly revealing surface features at high magnification.Finally, we couple a neon ion beam with the HIM setup allowing us tomill away surface membranes and provide us with unprecedentedaccess to sub-surface structures. The viability of this approach cen-ters on the sputter rates of neon being four times lower than that ofgallium19. As such, the neon ion beam is ideally suited for the pre-cision milling of very small and delicate biological structures. Inaddition, it causes far less discernable sample damage, or metal iondeposition, as is sometimes the case when using liquid metal ionsources such as gallium37,38.

The resolution attained using HIM imaging is far greater than thatobtained from a traditional FE-SEM approach. This higher resolu-tion necessitates the development of new protocols for sample pre-paration, particularly to mitigate the possibility of fixation anddrying artifacts. Cryo-immobilization offers a proven and reliablealternative for the current fixation and dehydration approaches.We suggest that combining this cryo-based sample preparation para-digm with HIM will allow routine imaging with little or no artifacts.

In conclusion we believe that this novel imaging method enablingthe observation of completely unmodified structures on the nan-ometer scale will revolutionize our ability to interrogate both surfaceand sub-surface structures and provide deep scientific insights intothe nanoscale architecture of biological specimens and soft materials.

MethodsSamples were grown/cultured and subsequently fixed and critical point/freeze driedusing the following methods. After the drying stage, the samples were carefullyremoved and adhered to double-sided carbon tabs on aluminum stubs and stored in adesiccator. For SEM imaging, the Arabidopsis thaliana, HeLa cells and Pristionchuspacificus samples were imaged on a Zeiss Sigma VP FE-SEM (Carl Zeiss, Cambridge,UK) and the Acidovorax sp. samples were imaged on a Zeiss Crossbeam 1540XB (CarlZeiss, Oberkochen, Germany) using the parameters mentioned above. All sampleswere subsequently imaged on a Zeiss Orion PLUS HIM (Carl Zeiss, Peabody, MA).

Arabidopsis thaliana. Growth. Wild type Arabidopsis were grown in soil to stage 6.Plants were trimmed by carefully pinching off regions of interest using two pairs of #5forceps and the subsequent pieces were placed into ice-cold 100% EM grade ethanolfor fixation.

Critical point drying. The ethanol fixed plant pieces were placed into a Teflon carrierfor separation and into an automated critical point drier (Leica EM CPD300, LeicaVienna) which was set to perform 25 exchange cycles of CO2 at medium speed and20% stirring. All additional fill, heating, and venting steps were performed at mediumspeed as well. After drying, the samples were carefully removed and adhered todouble-sided carbon tabs on aluminum stubs and stored in a desiccator.

HeLa cells. Coverslip preparation. Cleaned 22 3 22 mm no. 1.5 coverslips were coatedwith a 13 solution of poly-L lysine for 15 minutes and air-dried at room temperature.Coated coverslips were then sterilized by UV light (NuAire cell culture cabinet) for 15minutes on each side and stored until use.

Cell culture. HeLa cells were cultured in DMEM (Sigma-Aldrich) supplemented with10% fetal bovine serum, 1.0% l-glutamine with 0.018% NaHCO3 and penicillin/streptomycin in 5% CO2 and 95% air in a Thermo Scientific Forma Series II WaterJacketed CO2 cell culture incubator at 37uC. Cells were allowed to grow for at least 36hours from the previous passage until reaching 60–80% confluency before passageagain. Cells were washed twice with 2–5 mls of room temperature 13 DPBS and thetrypsinized with 1–2 mls of 0.05% Trypsin-EDTA (Life Technologies) for 5–10minutes at 37uC. Trypsin was neutralized with 2–4 ml of 37uC complete media andcollected in a 15 ml conical vial. 80,000–120,000 cells (per well) were plated in 6-wellplates containing 3 mls of complete media and a single poly-L lysine coated coverslip.Cells were allowed to grow for 36–48 hours to ensure that the cells enter the expo-nential growth phase prior to fixation. Shorter periods of growth or over confluencyyielded a much lower percentage of mitotic cells upon fixation.

Cell fixation. Plated cells were gently rinsed with 2 mls of 37uC DPBS and aspiratedoff prior to fixation. 3–5 mls of freshly prepared 37uC fixation solution containing 4%paraformaldehyde and 2% glutaraldehyde in DPBS was applied for 30 minutes at37uC and swapped with fresh 37uC fixative to continue fixing for 1.5 hours at 37uC.Once the cells were fixed they washed briefly in DI water and were either criticallypoint dried or were stored at 4uC in 1% paraformaldehyde and 2% glutaraldehyde inDPBS.

Critical point drying. Fixed coverslips were placed into an automated critical pointdrier (Leica EM CPD300, Leica Vienna) which was set to perform 25 exchange cyclesof CO2 at medium speed and 20% stirring. All additional fill, heating, and ventingsteps were performed at medium speed as well. After drying, the samples werecarefully removed and adhered to double-sided carbon tabs on aluminum stubs andstored in a desiccator.

Iron-oxidizing bacteria (BoFeN1). Cell culture. Cultures of the nitrate-reducingAcidovorax sp. strain BoFeN1 were grown under anoxic conditions as describedpreviously33. In brief, cells were grown in anoxic freshwater medium with initialconcentrations of 10 mM Fe21 and 10 mM nitrate and 5 mM acetate. Samples of thecell suspension were taken with syringes after 5 to 6 days of incubation when Fe(II) iscompletely oxidized39.

Critical point drying. 1 mL of a centrifuged aliquot of the cell suspension were treatedwith 2.5% glutaraldehyde at 4uC for 1 hour. The samples were rinsed twice withdeionized water and deposited onto 10 mm glass cover slips that were previouslycoated with a 8 nm Pt-layer and poly-l-lysine. After 20 minutes to ensure binding ofcell-mineral aggregates to the poly-l-lysine layer, the samples were rinsed with de-ionized water und subsequently dehydrated in a series of ethanol dilutions (30%, 70%,95% and 33 100% dried on molecular sieve) with an equilibration step of 10 minuteseach. The samples were then placed in a critical point dryer (Polaron E3000), whereinthe ethanol was replaced by liquid CO2 at 1uC. After flushing of 3 3 200 g CO2 with20 minute equilibration periods after each step the temperature was graduallyincreased to 38uC while the pressure was monitored and adjusted to reach and remainat 84 bars. This ensured bypassing the critical point when the pressure was releasedfrom 84 bars to atmospheric pressure. Half of the samples were then sputter-coatedwith a 4 nm platinum layer using a BALZERS-UNION MED 010 (BAL-TEC,Liechtenstein) sputter coater. After reaching a vacuum of 8*1026 mbar, the sputteringprocess at was started in 2*1022 mbar of Ar.

Rapid freezing followed by freeze-drying. To be able to compare samples treated afterthe standard sample preparation protocol of CPD with a more natural state of the cell-mineral aggregates, we decided to use a physical fixation (rapid-freezing) instead ofchemical fixation, followed by freeze-drying under high vacuum. Since the cell-mineral aggregates of BoFeN1 are within the mm size ranges, high-pressure freezingwas not required to avoid the formation of ice crystals, but plunge-freezing in liquidpropane was sufficient. Therefore, droplets of the cell suspension were deposited ontoformvar-coated 300 mesh TEM-grids. They were blotted with filter paper andimmediately shot into liquid propane at liquid-nitrogen (LN2) temperature(,2196uC). The grids were stored in LN2 until freeze-drying. Freeze-drying wasdone on the cooling stage of a BAL-TEC BA 500 high vacuum coater. The grids wereplaced on the cooling stage at 2120uC. Thermal conduction between the sample andthe stage was ensured by a small droplet of liquid propane, which also prevented iceformation on the stage during sample transfer. The stage was gradually heated up to282uC, while maintaining the pressure in the recipient below 5*1025 mbar. Theprogress of freeze-drying was followed visually. Upon complete disappearance of theice after approximately 35–45 minutes, the stage was gradually heated up to 125uCbefore the chamber was vented and the samples could be removed. Half of the sampleswere sputter-coated with a 4 nm Pt layer as described previously.

Predator nematode - pristionchus pacificus. Worm culture. Pristionchus pacificusspecimens (strain identification: rlh24) were raised on agar plates coated with E. coli(OP50)40. At the young adult stage, worms were rinsed 33 in saline buffer (M9) andtransferred to an uncoated agar plate. Worms then remained off food for 5 hours,providing sufficient time for P. pacificus to consume residual E. coli adherent to their

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exterior cuticle. This process is critical to produce a clean imaging sample. Thepredators were then fixed by placing them into a solution of ice-cold 100% EM gradeethanol.

Critical point drying. The ethanol fixed worms were placed into a 30 mm porous pot(Electron Microscopy Sciences (EMS) cat# 70187-20) and into an automated criticalpoint drier (Leica EM CPD300, Leica Vienna) which was set to perform 25 exchangecycles of CO2 at medium speed and 20% stirring. All additional fill, heating, andventing steps were performed at medium speed as well. After drying, the samples werecarefully removed and adhered to double-sided carbon tabs on aluminum stubs andstored in a desiccator.

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AcknowledgmentsWe gratefully acknowledge financial support from the Waitt Advanced BiophotonicsCenter endowment (J.M.K., M.S.J. and J.A.J.F.), NCI P30 Cancer Center Support GrantCA014195-39 (Y.J.S. and J.A.J.F.), NINDS P30 Neuroscience Center Core GrantNS072031-01A1 (M.S.J. and J.A.J.F.) and DFG Emmy Noether grant OB 362/1-1 (F.Z. andM.O.). We would also like to thank the Ion Microscopy Innovation Center at Carl ZeissMicroscopy (Peabody, MA) for providing access to an Orion PLUS helium ion microscope,Carl Procko from the Chory lab at the Salk Institute for providing wild-type ArabidopsisThaliana for HIM imaging experiments and Kat Kearny for critical reading of themanuscript and for helpful suggestions regarding organization and formatting.

Author contributionsM.S.J., J.M.K. and Y.J.S. were responsible for the preparation of the plant and cell culturesamples and M.S.J. and C.H. undertook the plant, nematode and cell HIM imagingexperiments. C.H. and D.F. undertook the neon cutting of nematode experiments. F.Z.,M.O. and C.B. were responsible for the preparation of the bacterial samples and the bacterialHIM imaging. Worm samples were prepared by K.C. and S.H.C. Study was conceived byB.G. and J.A.J.F., M.S.J. and J.A.J.F. prepared the figures and wrote the manuscript withcontributions from M.O., C.B., L.A.S. and B.G.

Additional informationSupplementary information accompanies this paper at http://www.nature.com/scientificreports

Competing financial interests: The author(s) declare that Chuong Huynh, David Ferranti,Lewis Stern and Bernhard Goetze are employed by the manufacturer of the commercialOrion PLUS Helium Ion Microscope (Carl Zeiss Microscopy, LLC). No other competingfinancial interests apply.

How to cite this article: Joens, M.S. et al. Helium Ion Microscopy (HIM) for the imaging ofbiological samples at sub-nanometer resolution. Sci. Rep. 3, 3514; DOI:10.1038/srep03514(2013).

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